Expansion Microscopy on Saccharomyces cerevisiae

The unicellular eukaryote Saccharomyces cerevisiae is an invaluable resource for the study of basic eukaryotic cellular and molecular processes. However, its small size compared to other eukaryotic organisms the study of subcellular structures is challenging. Expansion microscopy (ExM) holds great potential to study the intracellular architecture of yeast, especially when paired with pan-labelling techniques visualising the full protein content inside cells. ExM allows to increase imaging resolution by physically enlarging a fixed sample that is embedded and cross-linked to a swellable gel followed by isotropic expansion in water. The cell wall present in fungi – including yeast – and Gram-positive bacteria is a resilient structure that resists denaturation and conventional digestion processes usually used in ExM protocols, resulting in uneven expansion. Thus, the digestion of the cell wall while maintaining the structure of the resulting protoplasts is a crucial step to ensure isotropic expansion. For this reason, specific experimental strategies are needed, and only a few protocols are currently available. We have developed a modified ExM protocol for S. cerevisiae , with 4x expansion factor, which allows the visualisation of the ultrastructure of the cells. Here, we describe the experimental procedure in detail, focusing on the most critical steps required to achieve isotropic expansion for ExM of S. cerevisiae .


Figure 1. Yeast expansion microscopy overview a) Scheme of expansion microscopy protocol workflow. b) Maximum intensity projection of DIC/DAPI composite image of yeast, acquired with a Nikon HCS using a 100x/1.45 oil objective. c)
Maximum intensity projection of fluorescence image of post-expanded yeast, acquired with a Zeiss LSM 980 using a 40x/1.1 water objective and airyscan detection. Signals of DAPI and DIC/NHS-Ester are represented in magenta and grey, respectively. Scale bars: 5 µm. Samples were assessed by manual segmentation and measurement of the area and circularity between expanded (n=13) and non-expanded (n=27) specimens. The average expansion factor across all experiments was 3.52 ± 0.42 (mean ± standard deviation) while the circularity was retained within the level 20% of the population's standard deviation.

Description
The unicellular eukaryote S. cerevisiae represents an invaluable resource for the study of basic eukaryotic cellular and molecular processes. The combination of a high genetic amenability, numerous genetic tools, and vast genomic resources makes it one of the most versatile model organisms, employed in a wide range of basic research disciplines. However, its small size compared to other eukaryotic organisms has limited its use for the study of sub-cellular structures. The diameter of unbudded yeast cells ranges approximately between 4 μm in haploids and 6 μm in diploids (Milo and Phillips 2015), complicating conventional diffraction-limited light microscopy approaches.
The last decade witnessed the rise of super-resolution (SR) techniques that enable sub-diffraction resolution fluorescence imaging of cellular structures. Expansion microscopy (ExM), for instance, allows for increasing imaging resolution by physically enlarging a fixed sample that is embedded and cross-linked to a swellable gel and is then expanded isotropically in water (Wassie et al. 2019). By doing so, fluorescently-labelled structures that previously would be too small or too close together to be distinguished can be resolved in 3 dimensions. The increased z-resolution can be exploited by z-sectioning, using confocal or light-sheet microscopy. In addition, ExM can also be used in combination with other super-resolution methods, such as airyscan confocal microscopy and Structured Illumination Microscopy (SIM), further enhancing resolution. ExM holds great potential to study the cellular structures of yeast, especially when paired with pan-labelling techniques to visualise total protein content inside cells. ExM protocols present a robust tool to study cellular organelles, obtaining superresolution results with the advantage of employing inexpensive and straightforward equipment compared to other techniques that require more complex and expensive setups, such as optical SR approaches and electron microscopy.
ExM can be applied in various organisms and cellular models, offering outstanding outcomes with expansion factors ranging from 4x to 20x (Faulkner et al. 2020, Truckenbrodt et al. 2018). However, certain cellular organelles are resistant to these processes, such as the cell wall of fungi and Gram-positive bacteria. These organisms present a cell wall composed of peptidoglycans or glycoproteins that are highly resistant to denaturation and conventional digestion processes. The digestion of the cell wall and the maintenance of the structure of the resulting protoplasts are crucial steps to achieve isotropic expansion. For this reason, specific experimental strategies are needed, and only a few protocols are currently available (Götz et al. 2020). In particular, only one study describes an ExM protocol applied to S. cerevisiae, showing different yeast structures in high resolution (Chen et al. 2021).
Here we describe an adapted and extended ExM protocol for S. cerevisiae ( Figure 1) with a final expansion factor of ~3.5x. The major difference regarding to the protocol of Chen et al., is the staining strategy. While the previously published protocol is based on immunolabeling to visualise and super-resolve specific target proteins, here, samples are stained with DAPI and the pan-labelling reagent NHS-Ester BODIPY FL. This allows to provide an overview of the ultrastructural context of the whole cells. However, both protocols complement each other and could be combined, in principle. Here, we have compiled detailed step-by-step instructions for ExM to visualise the ultrastructure of S. cerevisiae, supported by supplementary video documentation (Supplementary Video 1).
Briefly, the first day begins with the preparation and fixation of yeast cells, which includes the digestion of the cell wall with zymolyase. The generated protoplasts are mounted on the coverslip, and linking takes place overnight, during which linker molecules will attach to the sample's proteins, acting as anchors for the next step. On the second day, the first step is gelation, where the sample is embedded in a monomer solution that polymerases and forms a matrix to which the anchor molecules will be connected. This is followed by the denaturation step, where the sample structures are disintegrated. The samples are then left expanding overnight in water, swelling the polymerised gel, which will push the anchored molecules apart in an isotropic manner. Finally, on the third day, staining with DAPI and NHS-Ester takes place. Samples are left again expanding overnight, and they can be imaged from the next day onwards.
Assessing the expansion factor and specimen cellular integrity following sample preparation is a critical step that requires consideration. During sample expansion, the denaturation/digestion steps facilitate isotropic expansion of the sample but different organelle structures might possess specific requirements that need to be identified during the experimental planning phase of the experiment. The simplest solution to assess specimen integrity and expansion factor is to acquire microscopy data of the same field of view and specimen before and after expansion. This process will allow the user to detect immediate changes in the sample but carries the complication of being difficult to execute since finding the same sample, particularly in yeast, can be difficult and time-consuming. Other options, such as the one included in this protocol, image multiple expanded and non-expanded cells and compare their features (area and circularity) as means to assess expansion (average circularity 0.892 ± 0.063 on expanded samples (mean ± standard deviation, n=13) and 0.906 ± 0.035 on non-expanded ones (n=27)). Individual organelle expansion should be obtained via specific labelling of the structure and determining before and after expansion sizes and compare it to the expansion factor of the whole cell.

Preparation of stock solutions:
Sorbitol buffer: Linking solution: • 0.1 mg/mL acryloyl X-SE in PBS. Store in aliquots at -20 °C.
YPD Media: 1. Dissolve 5 g of yeast extract and 10 g of peptone in 375 mL of ddH 2 O by manually stirring the flask and autoclave the solution.

Coverslip cleaning
Notes: i) This procedure should be performed under a fume hood, and one should use safety gloves when handling chloroform.
ii) The chloroform and NaOH solutions can be reused. iii) The preparation of NaOH is an exothermic reaction, thus the NaOH stock solution should be prepared on an ice bath. iv) Other coverslip cleaning protocols are available, such as a chloroformfree protocol (Pereira et al. 2015).
1. Arrange single coverslips in a coverslip rack.
2. Place a glass container in an ultrasonic bath and fill the bath with water.
3. Place the rack in the glass container.
4. Fill the glass container with chloroform and sonicate for 1 hour.

5.
Remove the rack with a tweezers/forceps and place it in an empty glass container. The coverslips will dry in a few minutes as the chloroform evaporates quickly.
6. Pour the remaining chloroform back into the bottle through a funnel.
7. Reinstall the rack in the glass container in the ultrasonic bath.
8. Fill the glass container with a 5 M NaOH solution and sonicate for 1 hour.
9. Pass the coverslips into a beaker with ddH 2 O and transfer the NaOH solution back into the bottle.
10. Wash the coverslips three times in a volume of 500 mL fresh ddH 2 O.
11. Pre-dry the coverslips with a nitrogen or filtered air gun and then place them in a drying cabinet until they are completely dried.
12. Store in 100 % ethanol in a glass Petri dish.
2. Place the coverslips in a clean plastic plate, for example, in a 6-well plate.
3. Add 300 µL of 0.1 mg/mL poly-L-lysine on each coverslip and incubate at room temperature (RT) for 10 minutes.
4. Add water to each well until the coverslip is covered and incubate at RT for 5 minutes.
6. Store coverslips on a rack at 4 °C until needed for cell/gel mounting.

Yeast cells cultivation and fixation
The Saccharomyces cerevisiae BY4741 strain (MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0) was used in this protocol.
1. Culture yeast cells in the conditions required to assess the relevant biological question. (In our case, we incubate yeast cells in fresh YPD media supplemented with adenine and tryptophan at 30 °C and 220 rpm of agitation until the culture growth reaches the exponential phase).
2. Count yeast cells with a hemocytometer and calculate the volume required to collect 6x10 7 cells.
3. Centrifuge cells at 4,000 rpm for 2 minutes in a Falcon tube. Linking 1. Remove PBS, add 1 mL of 0.1 mg/mL acryloyl X-SE solution in PBS to each well and incubate for 12 hours (overnight) at RT.

Day 2
Gelation Note: Proceed quickly through steps 3 to 5, as after the gelation solution is prepared, it will polymerise in minutes. For this reason, it is recommended to proceed with these steps with a maximum of four samples each time. Reagents and slides are maintained on ice to prevent premature polymerisation.
1. Cool the reagents (monomer solution, TEMED, and APS) and the gelation chamber slides (see supplementary figure 1) on ice for at least 10 minutes.
2. Remove the excess solution from the coverslip with a lint-free wipe and place it next to the slide on which it will be mounted (Note: the coverslip side with the sample should be facing up).
3. Prepare the gelation solution: Add first the TEMED to the monomer solution, then APS and vortex. (Note: it is important to follow this order, as when APS is added the polymerisation reaction starts).